Abstract

Cell division is essential for tumor development and progression. Methylation-mediated silencing caused by aberrant de novo methylation of CpG islands located in the promoter regions of growth regulatory genes occurs frequently in human cancers. We investigated the relationship between cell division and de novo methylation to determine whether de novo methylation can occur in the absence of cell division in cancer cells. We treated T24 bladder carcinoma cells with 5-Aza-2′-deoxycytidine to induce a transient demethylation and then compared the timing and kinetics of remethylation of the p16 gene locus under conditions of either G0-G1 growth arrest induced by serum starvation and confluence or continuous cell proliferation in complete medium. Variable levels of remethylation were detected in CpG poor regions of DNA, as well as repetitive DNA elements in the absence of cell division, yet no remethylation occurred at CpG islands under these conditions. This correlated with continuous expression of p16 protein in these cells. DNA methyltransferase (DNMT)1 and DNMT3b3 proteins were undetectable in 5-Aza-2′-deoxycytidine-treated and untreated nondividing cells, and their mRNA transcripts were down-regulated in these cells. Although DNMT3a mRNA levels were also reduced, they recovered to original levels in nondividing cells after drug treatment. Our results suggest that cell division is required for de novo methylation of CpG islands and that DNMT3a may play a role in methylating CpG poor regions or repetitive DNA elements outside of the S phase of the cell cycle.

INTRODUCTION

De novo DNA methylation occurs early during normal mammalian embryonic development by the addition of a methyl group to the 5′ position of a previously unmethylated cytosine in the context of a CpG dinucleotide. The number of CpG dinucleotides in the genome is underrepresented by a factor of five
(1)
. Most cytosine methylation occurs in CpG poor regions of DNA, as well as in repetitive elements
(2)
. More than 50% of the remaining unmethylated CpG sites reside in CpG-rich DNA regions known as CpG islands, which are often present within or near the promoters of genes
(3, 4)
. In sharp contrast, CpG islands located in promoter regions of genes on the inactive X chromosome are not protected against de novo methylation
(4)
. An inverse correlation has been shown to exist between DNA methylation status at promoter CpG islands and the level of gene expression attesting for the role of DNA methylation as a transcriptional repression mechanism
(5)
.

One of the most frequent alterations present in human cancer cells is the aberrant de novo methylation of CpG islands
(6, 7)
. Hypermethylation of promoter-associated CpG islands can lead to loss of gene expression and gene silencing
(7)
. In cancer cells, this may represent an alternative mechanism to deletions or mutations to inactivate tumor suppressor genes. One possible explanation as to why cancer cells fail to maintain the CpG islands in an unmethylated state is that such cells may harbor defects in their DNA methylation machinery. There are three known DNMTs
4
described in mammalian cells: DNMT1, 3a, and 3b. The task of de novo methylation of CpG sites during embryogenesis appears to be shared by DNMT3a and 3b
(8)
, although both enzymes have also been shown to equally methylate both unmethylated and hemimethylated DNA in vitro with similar efficiency
(9, 10)
. Conversely, DNMT1 seems to be the enzyme primarily responsible for maintenance methylation, as it has a preference for hemimethylated DNA in vitro. DNMT1 has also been shown to have a limited de novo activity in vitro(11)
but no detectable de novo activity in Drosophila(12)
. In addition, each individual methyltransferase appears to preferentially target specific regions of DNA. Sequence-specific DNA methylation defects have been described in patients with Immunodeficiency, Chromosomal Instability, and Facial Abnormalities syndrome
(13, 14)
, a rare hereditary disease caused by mutations in the DNMT3b gene
(8, 15, 16)
.

Several studies suggested that abnormal de novo methylation seen in cancer cells may be attributed to changes in the levels of expression of DNMTs. All DNMTs have been shown to be overexpressed at the RNA level to various degrees in several cancers
(17,
18,
19,
20)
. Overexpression of murine Dnmt1 was shown to induce transformation
(21)
, whereas inhibition of this enzyme by antisense constructs or treatment with methylation inhibitors prevented tumorigenesis
(21)
. Aberrant de novo methylation of endogenous CpG islands has been documented after overexpression of Dnmt1 in normal cells
(22)
.

We proposed recently that methylation alterations detected in some cancer cells could alternatively be attributed to improper expression of DNMTs during the cell cycle
(23)
. The mRNA levels of these enzymes were shown to be differentially regulated during the cell cycle and that changes in this regulation could distinguish normal cells from tumor cells. Additional studies have demonstrated de novo methylation of endogenous CpG islands after 3 weeks of growth arrest in normal cells
(24)
, and limited remethylation was shown to occur after DNA excision repair in arrested cells damaged with UV radiation
(25)
. Remethylation of CpG islands in cancer cells after treatment with a demethylating agent was shown to be independent of the rate of cell division
(26)
.

In this study, we investigated whether de novo methylation can occur in nondividing cancer cells. We treated a bladder cancer cell line with 5-Aza-CdR to induce a transient demethylation. We then monitored the kinetics of remethylation at specific CpG sequences within the p16 gene locus, as well as DNMT expression levels in cells that subsequently were either allowed to proliferate or were maintained in a nondividing state. Here we show that DNMT1 and DNMT3b3 mRNA transcripts remained down-regulated and their proteins undetectable in 5-Aza-CdR-treated and untreated nondividing cells. DNMT3a mRNA recovered to original levels in these cells. No remethylation of CpG islands occurred in nondividing cells, whereas variable degrees of remethylation were detected in the CpG poor regions and repetitive elements. These results suggest that cell division is required for de novo methylation of CpG islands in cancer cells.

5-Aza-CdR Treatments.

T24 cells were plated at 2 × 106 cells/100-mm dish and were treated with 1 × 10−6m 5-Aza-CdR 24 h later. The medium was then changed 24 h after drug treatment and every 3 days thereafter. At the 3rd day after 5-Aza-CdR treatment, cells were confluent. At this point, one half of the cells were split by trypsinization and were subsequently seeded and kept in a dividing state by maintenance in 10% FCS. The other cells were maintained in a confluent state (nondividing) grown in 0.1% FCS.

Flow Cytometry Analysis.

Cells (106) were pelleted and resuspended in 200 μl of PBS, fixed in 2 ml of ice-cold ethanol, and centrifuged again to remove the fixative. Cell pellets were resuspended in 1 ml of PBS containing 10 μg/ml propidium iodide. The fluorescence was measured on a Fluorescence Activated Cell Sorter Plus flow cytometer (Becton Dickinson, San Jose, CA). Population doublings were determined by measuring the total cell number at the times of seeding and collection using a Coulter Counter (Coulter Electronics, Hialeah, FL; model Zf).

BrdUrd Incorporation Assay.

T24 cells maintained in culture in either 10% FCS (dividing) or 0.1% FCS (nondividing) were incubated with 10−4m BrdUrd (Sigma Chemical Co., St. Louis, MO) for 90 min. Fixation and denaturation were performed as recommended by Becton Dickinson. The anti-BrdUrd antibody was obtained from Becton Dickinson, and antimouse-IgG-FITC antibody was obtained from Caltag Laboratories (Burlingame, CA). Indirect immunofluorescence staining of BrdUrd-incorporated cells and propidium iodide labeling was performed as recommended by Becton Dickinson. Cells were analyzed on a Calibur flow cytometer from Becton Dickinson at an excitation wavelength of 488 nm. Cell cycle distribution was determined using CellQuest software from Becton Dickinson.

Nucleic Acid Isolation.

DNA and RNA were collected and extracted from both dividing and nondividing cells at various time points throughout the experiment as described previously
(27)
. Total cellular RNA was purified as described in Bender et al., 1998
(28)
.

Bisulfite Treatment of Genomic DNA.

Genomic DNA (4 μg) was treated with 40 units of EcoRI (Roche, Indianapolis, IN) at 37°C for 16 h. DNA was incubated with 0.3 m sodium hydroxide for 20 min at 45°C, followed by the addition of 3.6 m sodium bisulfite (pH = 5.0) and 0.11 m hydroquinone for 16 h at 55°C. The reaction mixture was then purified with the Promega Wizard Mini-Prep Kit (Madison, WI) and desulphanated with 0.3 m sodium hydroxide for 20 min at 40°C. The DNA was then precipitated in three volumes of cold ethanol, dissolved in H2O, and stored at −20°C.

Bisulfite-specific PCR.

Bisulfite-converted DNA was PCR amplified using the primers shown in Table 1
⇓
. Each PCR mixture contained 100 μm deoxynucleotide triphosphates, 1 μm sense and antisense primers in 1 × Taq Buffer, and 1.25 units of Taq DNA polymerase (Sigma Chemical Co.) complexed with 1.25 units of Taq antibody (Clontech, Palo Alto, CA). Each PCR program was as follows: 95°C for 3 min, followed by 40 cycles of denaturation at 95°C for 1 min, annealing at the specific temperature listed in Table 1
⇓
for 45 s, and finally, a 45-s extension at 72°C. A final 10-min extension at 72°C completed each PCR program. PCR products were fractionated on 1% agarose gels, excised, and purified with the Qiagen DNA extraction kit (Valencia, CA) according to manufacturer’s recommendations. The purified DNA was redissolved in H2O and stored at −20°C.

Quantitation of DNA Methylation Levels by Ms-SNuPE Assay.

The Ms-SNuPE method for determining methylation levels was performed as described by Gonzalgo and Jones
(29)
. Dual reactions (10 μl each) were performed with purified bisulfite-treated PCR fragments of interest using the Ms-SNuPE primers in Table 1
⇓
in the presence of either 32P-dCTP or 32P-TTP with 0.05 units of Taq DNA polymerase complexed with 0.05 units of Taq antibody. The Ms-SNuPE reactions were performed for all regions as follows: 95°C for 2 min, 50°C for 2 min, and 72°C for 1 min. The incorporation of C identifies a methylated CpG dinucleotide, whereas the incorporation of T identifies an unmethylated CpG dinucleotide. Stop buffer [4 μl of formamide, 1 μm EDTA (pH = 8.0), 0.1% bromphenol blue, and 0.1% xylene cyanol] was added, the samples were denatured, and 1.5-μl aliquots were electrophoresed on 15% acrylamide sequencing gels. The gels were dried and analyzed using a Molecular Dynamics PhosphorImager 445 SI (Sunnyvale, CA). Each Ms-SNuPE primer is specific for a single CpG dinucleotide. The methylation status of regions containing multiple CpG dinucleotides
(1, 2, 3, 4, and 8)
represents the averaged methylation levels of three to four CpG sites. All CpG sites analyzed in a given region were similar in their methylation status. Because the methylation levels and remethylation kinetics of the CpG poor sites 1 and 2, as well as 5 and 6, were similar, the results for these regions were also combined in the final interpretation of the data.

Northern Blot Analysis.

Total RNA (10 μg) extracted from untreated or 5-Aza-CdR-treated T24 cells was fractionated on 1% formamide-agarose gels and transferred to nylon membranes. 32P-labeled cDNA probes specific for DNMT1, 3a, and 3b
(23)
were used for hybridization. Quantitation of signal intensities of Northern blots was achieved using a PhosphorImager.

Western Blot Analysis of p16 and DNMT Protein Levels.

Cells in a 100-mm dish were rinsed with two volumes of ice-cold PBS followed by the addition of 75 μl of radioimmunoprecipitation assay buffer (1 × PBS, 1% SDS, 0.5% NP40, and 0.5% sodium deoxycholate). The cells were scraped off the dish and placed on ice for 30 min. The mixture was then centrifuged at 13,000 rpm for 30 min at 4°C. The supernatant was removed and used for Western analysis. Approximately 20 μg of total protein extract were loaded onto 4–15% gradient Tris-HCl gels (Bio-Rad, Hercules, CA), electrophoresed in Tris-glycine-SDS running buffer, and transferred to a polyvinylidene difluoride membrane in Tris-glycine buffer overnight at 4°C. The membranes were hybridized with antibodies against DNMT1 (1:1000 dilution; New England Biolabs), DNMT3b (1:100 dilution; Santa Cruz Biotechnology), and p16 (1:200 dilution; Santa Cruz Biotechnology) in Tris-buffered saline-T buffer with 5% nonfat dry milk for 1 h at room temperature. The membranes were washed five times with Tris-buffered saline-T at room temperature. The membranes were then incubated with secondary antibodies as follows: antimouse-IgG-HRP (1:1000 dilution for p16; Santa Cruz Biotechnology), antirabbit-IgG-HRP (1:1000 dilution, for DNMT1; Santa Cruz Biotechnology), and antigoat IgG-HRP (1:1000 dilution; Santa Cruz Biotechnology) for 1 h at room temperature. The proteins were detected with the enhanced chemiluminescence detection kit (Amersham-Pharmacia) and by exposure to Kodak X-OMAT AR film.

RESULTS

The two CpG islands located in the p16 gene have been shown to become targets for aberrant de novo methylation in human cancers
(27)
. These CpG islands, located in the promoter and second exon, are separated by a CpG poor region of DNA in the first intron (Fig. 1)
⇓
. Additional CpG poor regions, as well as Alu-repetitive elements, are located upstream of the promoter. We first used the quantitative Ms-SNuPE assay to determine the methylation status of these regions in LD419 normal bladder embryonic fibroblasts (Fig. 1)
⇓
. The methylation levels of several CpG dinucleotides within CpG-rich regions can be determined simultaneously with this assay
(29)
. The CpG sites analyzed by Ms-SNuPE analysis in the promoter and exon2 of the p16 gene have been shown previously to reflect the methylation status of each of these CpG islands as a whole
(26, 30,
31,
32)
. The CpG islands (regions 4 and 8) displayed low levels of methylation (5–15%), whereas the CpG poor regions (regions 1–2 and 5–7) and the Alu repeat (region 3) had higher methylation values (30–65%). Similar results were found in normal colonic epithelium tissue samples (data not shown), suggesting that the pattern in fibroblasts was not because of a cell culture artifact. This analysis clearly illustrated the high extent of protection from DNA methylation of the two CpG islands in the normal cells when compared with the adjacent CpG sites located on a continuous 13-kb DNA segment. In addition, the methylation level in an Alu repeat of the p16 gene (47%) was lower than that of an Alu-repetitive element located in the p53 gene (95%; data not shown), showing that these repeats are methylated variably in the human genome.

Methylation levels of regions within the p16 gene locus in normal LD419 embryonic fibroblasts and T24 bladder cancer cells either before and after treatment with 5-Aza-CdR. Top, eight regions of various CpG densities were identified upstream and downstream of the p16 promoter region. Regions 1 and 2 are CpG poor sequences (three CpGs analyzed in region 1 and four CpGs in region 2), region 3 is an Alu-repetitive element (four CpGs analyzed), region 4 is a CpG island located at the 3′ end of the p16 promoter (three CpGs analyzed), and regions 5–7 consist of three CpG dinucleotides residing in intron 1 of p16, whereas region 8 is the CpG island of the second exon (three CpGs analyzed). Vertical arrows, the specific CpG sequences analyzed by Ms-SNuPE. Bottom, the methylation values for each region were determined by Ms-SNuPE in LD419 cells (▪), and in T24 cells, before (
) and after (□) treatment with 5-Aza-CdR. T24 cells were treated with the drug for 24 h and allowed to proliferate for 2 additional days in drug-free medium. Day 3 represents the time point of maximum demethylation after drug treatment.

In contrast to the fibroblasts, all eight regions analyzed, irrespective of their CpG content or sequence, were hypermethylated in the T24 bladder cancer cells (Fig. 1)
⇓
. A lower degree of hypermethylation was detected however in two of the intronic CpG sequences (regions 5 and 6) flanked by the two CpG islands in comparison with the other regions. Therefore, all of the CpG sites studied in T24 cells showed increased methylation when compared with the fibroblasts.

We next evaluated the demethylating effect of 5-Aza-CdR 3 days after treatment of T24 cells with 5-Aza-CdR (Fig. 1)
⇓
, when the maximum level of demethylation was observed
(26)
. All regions examined were comparably demethylated after treatment with 5-Aza-CdR. Thus, this drug induced a global demethylation that equally affected abnormally methylated CpG islands, CpG poor regions, or CpG sites associated with repetitive elements.

Kinetics of Remethylation in Dividing or Nondividing T24 Cells after 5-Aza-CdR Treatment.

Remethylation of CpG islands in T24 cells after 5-Aza-CdR treatment has been shown to occur slowly, in a sequence-specific manner, independent of the rate of cell division and shown to be the result of a de novo mechanism by single cell cloning experiments
(26, 30)
. Although this process may not recapitulate the de novo methylation events occurring during tumorigenesis, at present time, it represents the most effective way to study de novo methylation of CpG sites that are hypermethylated in human cancers. In this study, we determined whether remethylation after 5-Aza-CdR treatment could occur in the complete absence of cell division. T24 cells were treated with 5-Aza-CdR for 24 h, grown to confluence at day 3, and then either allowed to proliferate or maintain in an arrested state (G0-G1) by confluence and serum starvation (Fig. 2A)
⇓
. The cell proliferation status of the cells was monitored by flow cytometry analysis. The presence of the G2 and S phases on the DNA histograms confirmed that the cells were actively dividing in 10% serum, whereas the absence of these phases in serum-starved cells were indicative of G0-G1 arrest (Fig. 2A)
⇓
. Analysis of BrdUrd incorporation combined with propidium iodide staining indicated that only 2% of the nondividing cells were in S phase in comparison with 21% of the dividing cells at day 27 after 5-Aza-CdR treatment (Fig. 2B)
⇓
.

The experimental approach to determine the extent of DNA methylation and the proliferation status in dividing and nondividing cells. A, experimental approach. Logarithmically dividing T24 bladder cancer cells growing in complete medium supplemented with 10% FCS (▪) were treated with 3 μm 5-Aza-CdR for 24 h (□), after which time, the drug was removed, and the cells were grown to confluence by day 3. At this time, some of the cells were kept in a nondividing state by serum starvation (0.1% FCS) and confluence (
), and some of the cells were split and allowed to actively proliferate in complete (10% FCS) media (▪). The proliferative state of the cells was monitored by flow cytometry, and representative DNA histograms of dividing cells (containing both G0-G1 and G2-M peaks) or nondividing cells (G0-G1 peaks only) are indicated. At the time points listed (days 0, 3, 6, 13, and 20), DNA, RNA, and protein extracts were isolated for assessing the extent of DNA methylation by Ms-SNuPE and gene expression levels by Northern blot and Western analyses. B, cell cycle analysis evaluating the proliferation status of T24 cells after BrdUrd incorporation. T24 cells actively dividing or growth arrested by serum starvation and confluence were treated with BrdUrd for 90 min. Cells were labeled with propidium iodide (PI) and subjected to indirect immunofluorescence staining with an anti-BrdUrd primary antibody and an FITC-conjugated secondary antibody as described in “Materials and Methods.” The proportion of unlabeled (R2) or BrdUrd-labeled (R3) cells in S phase was determined by flow cytometry analysis.

The rate of remethylation of p16-specific genomic regions in dividing and nondividing cells treated with 5-Aza-CdR was determined by Ms-SNuPE as a function of the lowest level of methylation after treatment (Fig. 3)
⇓
. The CpG poor regions upstream of the promoter (regions 1–2), the Alu repeat (region 3), the two CpG islands (regions 4 and 8), and one CpG intron 1 sequence (region 7) became 60–80% remethylated by day 20 in dividing cells, whereas the other two CpG sequences in the first intron (region 5–6) showed no remethylation in dividing cells (Fig. 3)
⇓
.

Remethylation kinetics of the CpG poor regions, the Alu-repetitive element, and the CpG islands in the p16 gene locus by Ms-SNuPE. Each region (highlighted in the map) was assayed by Ms-SNuPE at the time points listed in Fig. 2
⇓
. The PCR and Ms-SNuPE primers and conditions are listed in Table 1
⇓
, and the arrows on the map represent the CpG sequences analyzed. The transcription start site for the p16 gene is indicated by the bent arrow on the map. The numbering and identification of the regions are the same as in Fig. 1
⇓
. Each graph is plotted as a percentage of remethylation as a function of the value of methylation for each region at day 3, the time of maximum demethylation for each region. The remethylation values of cells in the dividing state are indicated as ♦ and those in the nondividing state as □. Regions 1 and 2 consistently showed similar rates of remethylation, so the values from both of these regions were combined into one graph (1 + 2). The data for regions 5 and 6 were also combined in a similar manner (5 + 6). The values represent the average of three to eight experimental trials, and the error bars represent the SD of those values.

Invariably, no remethylation was detected in either CpG island in the absence of cell division after multiple experimental trials (Fig. 3)
⇓
. No >16% remethylation of the PAX6 exon 5 CpG island was detected in nondividing cells (data not shown). These results suggest that the absence of CpG island remethylation in nondividing cells was not specific for the p16 locus. Various degrees of remethylation (20–60%) were detected at some but not all CpG poor regions in nondividing cells, as well as in the Alu repeat (Fig. 3)
⇓
. However, this variability was not only region specific but also occurred at the same region during the course of multiple experimental trials. The fact that remethylation was detected in nondividing cells also may suggest that the mechanisms responsible for the methylation of these sequences may be different from those methylating the islands. Because of the high degree of variability in the levels of remethylation observed for the CpG poor regions in nondividing cells, the significance of this finding will require future additional experiments to be fully understood.

One possible explanation as to why CpG islands fail to become remethylated in the nondividing cells is that residual 5-Aza-CdR may still be present in the genomic DNA after treatment by the time the cells reached confluence. This may hinder the remethylation process because cell division is required for removal of 5-Aza-CdR from genomic DNA. We repeated the treatment and then allowed the cells to divide for 3 additional days before reaching confluence. Again, no CpG island remethylation was observed under these conditions (data not shown).

Analysis of DNMT and p16 Expression Levels during Remethylation after 5-Aza-CdR Treatment.

We determined the effects of cell division on the expression levels of DNMTs and p16 during the remethylation process in T24 cells. The mRNA levels of DNMT1, 3a, and 3b3 (the only 3b transcript expressed in T24 cells
5
; Ref.
19
) were measured by Northern analysis at various time points in cells untreated or treated with 5-Aza-CdR (Fig. 4A)
⇓
. In the absence of this drug, all DNMT transcripts were down-regulated when cells were confluent, with the DNMT1 levels being the most reduced. The mRNA levels remained down-regulated while the cells were under growth arrest (Fig. 4B)
⇓
. After drug treatment, all DNMT transcripts were also down-regulated in nondividing cells (Fig. 4B)
⇓
. Although DNMT1 mRNA levels remained low and DNMT3b3 levels continued to decrease, DNMT3a mRNA rebounded to original levels by day 17 postconfluence (day 20 after treatment).

DNMT expression levels in nondividing-treated and untreated T24 cells. A, Northern blot image of DNMT mRNA expression levels in nondividing-treated and untreated conditions. Total RNA from T24 cells either untreated or treated with 5-Aza-CdR as described in Fig. 2
⇓
was used for Northern blot analysis. RNA fractionated by electrophoresis was transferred to nylon membranes, which were probed for DNMT1, 3a, and 3b3 presence. B, PhosphorImager quantitations of the Northern blots shown in A. The expression levels of DNMT1 (•), DNMT3a (▪), and DNMT3b3 (▴) were normalized to the levels of expression before treatment (log phase) for each individual enzyme and to the levels of 28S RNA quantitated from an ethidium bromide-stained agarose gel. Days −3 and 0 represent the time of treatment and achievement of confluence, respectively. C, Western blot analyses of DNMT1, DNMT3b3, and p16 protein levels in untreated cells and cells treated with 5-Aza-CdR. Cell lysates were obtained from treated and untreated T24 cells as described in Fig. 2
⇓
and were analyzed by Western blot analysis with specific antibodies for DNMT1, DNMT3b, and p16 proteins. The molecular weights of each protein are indicated.

Western blot analysis using antibodies specific for DNMT1 and 3b indicated that these proteins were not present in treated and untreated nondividing cells (Fig. 4C)
⇓
. DNMT1 and 3b3 protein levels fully rebounded in dividing cells by day 6 after release from confluence in the untreated cells and by day 13 after 5-Aza-CdR treatment. The delay in appearance of these proteins under treated conditions may be because of the decreased rate of cell division between days 6 and 13 (data not shown). These results indicated that the activity of these two enzymes may be restricted mostly to the S phase of the cell cycle, as their protein expression levels were significantly reduced outside of S phase. The status of DNMT3a protein levels could not be assessed because reliable antibodies against DNMT3a protein are not available commercially. However, the up-regulation of DNMT3a mRNA in drug-treated nondividing cells suggested that this enzyme may function in de novo methylation of CpG poor regions and repetitive elements in nondividing cells. However, the absence of remethylation of CpG islands in nondividing cells (Fig. 3)
⇓
provided evidence that although DNMT3a mRNA was present at this stage in the cell cycle, DNMT3a protein may not be active on these substrates.

We assessed the demethylating effect of 5-Aza-CdR on the p16 promoter by evaluating the levels of expression of p16 protein (Fig. 4C)
⇓
. No p16 protein was detected in untreated cells. In the treated cells, p16 protein was expressed by day 3. Its expression was maintained in the nondividing cells but was almost completely lost by day 13 in the dividing cells. Because de novo methylation of the promoter CpG island results in the down-regulation of p16 expression
(30)
, the continued expression of p16 protein in nondividing cells provided additional evidence that the promoter became methylated only in the dividing cells.

DISCUSSION

Abnormal de novo methylation of CpG islands occurs frequently in human cancers. In this study, we analyzed the methylation status of 20 CpG sequences in a 13-kb-long continuous segment of DNA harboring the p16 gene. The entire region was hypermethylated extensively in a bladder cancer cell line compared with normal bladder fibroblast cells. The two CpG islands were the most affected, because they were unmethylated in the normal fibroblasts but methylated extensively in the tumor cells. However, the CpG poor regions and an Alu-repetitive element were also hypermethylated when compared with normal fibroblast cells. These results suggest that CpG islands are not the only targets for de novo methylation in human cancers.

We next determined whether de novo methylation can occur in the absence of cell division for the regions described above. No remethylation was detected in nondividing cells in either p16 CpG island. The continuous expression of p16 protein in these cells provided additional evidence that the CpG islands remained demethylated under these conditions. Because the link between hypermethylation of the promoter and expression of the p16 gene has been established
(30)
, one possible explanation is that transcription of the p16 gene may interfere with the remethylation process. However, remethylation of the 3′ terminus of intron 1 detected in nondividing cells suggests that remethylation can occur on a transcribed gene. Similarly, a previous study from our laboratory
(26)
showed that transcription through a CpG island does not inhibit the remethylation process.

Our finding that de novo methylation of 3 CpG islands, including those located in p16 and one CpG island in PAX6 exon 5, did not occur in nondividing cells suggests that this may represent a global phenomenon. This is not in agreement with a previous study by Pieper et al.(24)
, who showed evidence of CpG island methylation in nondividing cells. We do not have a clear explanation for this discrepancy at present. However, the difference in the results between the two studies may be attributable to the use of different human cell types (epithelial bladder cancer cells, in the present study, compared with normal embryonic lung fibroblasts).

With the exception of two CpG sequences located in intron 1, which were not remethylated in either dividing or nondividing cells, all of the regions analyzed became remethylated in dividing cells. Various degrees of remethylation were detected for all CpG poor regions and the Alu sequence in nondividing cells. With the exception of regions 5 and 6, lower rates of remethylation were observed by day 20 in nondividing cells when compared with the dividing cells. This is in agreement with the findings of Kastan et al.(25)
, who showed that arrested cells damaged by UV radiation are capable of limited remethylation during DNA excision/repair process. Because a small percentage (2%) of nondividing cells escaped growth arrest as measured by BrdUrd incorporation, it is possible that this may have contributed to the remethylation of CpG poor and Alu regions observed in these cells.

The differences in the rates of remethylation in nondividing cells between CpG islands and the non-CpG island regions suggest that these sequences may be targeted by different DNMTs. Alternatively, this phenomenon may also be attributable to differences in chromatin structure at the different p16 gene regions, because our methylation analyses involved almost 13 kb of genomic DNA. Our analysis of the expression levels of DNMTs showed that DNMT1 and 3b3 mRNA transcripts and proteins were reduced in nondividing cells after 5-Aza-CdR treatment. This was in contrast with the levels of DNMT3a mRNA transcripts, which accumulated after drug treatment in these cells. Although DNMT3a protein levels could not be determined, the increase in its mRNA expression after day 10 in growth-arrested cells after drug treatment suggested that this enzyme may be responsible for de novo methylation of sequences, such as CpG poor and Alu-repetitive elements outside the S phase of the cell cycle. The incomplete remethylation of these regions may also suggest that DNMT3a, along with DNMT1 and/or 3b3, is required for their de novo methylation. Such cooperativity between DNMTs was also shown recently to be required for maintenance methylation of repetitive elements in mouse embryonic stem cells
(33)
.

DNMT3a may not be responsible, however, for the methylation of CpG islands when cells are nondividing, but it is possible that this enzyme can act on these sequences during the S phase, either by itself or together with other DNMTs that are only active during this phase of the cell cycle. However, the correlation between the absence of DNMT1 and 3b3 proteins and the lack of remethylation of the p16 CpG islands in nondividing cells suggests that these DNMTs may be responsible for their de novo methylation. Consistent with this hypothesis is the finding that CpG islands located on the inactive chromosome X are hypomethylated abnormally in patients with Immunodeficiency, Chromosomal Instability, and Facial Abnormalities syndrome that lack normal DNMT3b function
(14, 34)
.

The most likely explanation for the failure of CpG islands to become remethylated in nondividing cells is that de novo methylation of the CpG islands must occur only in cells that are dividing. This may explain why aberrant CpG island hypermethylation occurs in cancer cells in which loss of cell cycle control regulation results in unrestrained cellular proliferation. Epidemiological, as well as molecular, genetic studies suggest that cell division is essential for the genesis and progression of human cancers
(35)
. A variety of physical, chemical, infectious, and hormonal factors have been shown to contribute to the neoplastic process by stimulating cellular proliferation. Molecular genetic abnormalities, such as mutations, recombinations, or translocations, are believed to be clonally selected during the process of cell division. Loss of telomere ends leading to chromosomal rearrangements, as well as the possible reactivation of telomerase, also depends on multiple rounds of cell division
(36)
. We propose that similarly to the genetic errors, epigenetic changes in cancer cells, such as abnormal de novo methylation of CpG islands, may also require cell division during cancer development. This hypothesis is also supported by previous studies documenting de novo methylation in cultured senescent cells
(37,
38,
39)
, as well as age-related de novo methylation in normal colonic epithelium of elderly individuals
(40)
that may have undergone numerous rounds of cell division. Similarly, accelerated age-related CpG island methylation has been shown to occur in younger patients with ulcerative colitis, a chronic inflammatory disorder characterized by increased cell turnover and increased risk of colon cancer
(41)
. Our study suggests that nondividing cells are not targets for de novo methylation at these critical regions, whereas proliferating cells may be at higher risk for developing such changes.

Acknowledgments

We thank the members of Peter Jones’ laboratory for helpful discussions. We also thank Dr. Louis Dubeau for critically reviewing this manuscript and for helpful discussions.

Footnotes

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.